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Is it known if factors such as CD9 or CD81 are specific for exosomal origin?

Currently there is no consensus about a general exosome marker. The current recommendation from the research community is to combine detection of many membrane-bound or membrane-associated proteins in order to verify the presence of membranes. Our own experience is that targets CD63 and CD81 are found in many different exosome preparations, as is CD9. However, we detected at least 2 different cell lines releasing exosomes which were CD9 negative (Jurkat cells and several B-cell lymphoma cells, Oksvold et al 2014). Equally important is to document the absence of contaminating vesicles by staining for markers known to be expressed in compartments such as ER, Golgi or nucleus. It is also important to characterize the host cell in terms of the same markers.

Is it possible to quantify the number of exosomes by analyzing labeled exosomes bound to Dynabeads products using FACS?

Yes. The way the Dynabeads magnetic beads are designed and produced for cell isolation, protein isolation, nucleic isolation, T-cell activation and expansion (and now also exosome capture and isolation) helps ensure a high degree of reproducibility. All the beads are the same size and have a surface chemistry that provides very low unspecific binding and high signal/noise ratio. With this in mind, isolating exosomes using Dynabeads (paying attention to the bead concentration and capture volume) will enable constant binding kinetics. This setup will not give you the exact number of exosomes on the bead surface, but it will provide increased signal as a result of increased input in a linear way. Therefore, the beads can be used to measure exosomes, e.g., to monitor the exosome release efficiency from cells.

Electron microscopy must be used to estimate the number of exosomes per bead, i.e., isolate exosomes with Dynabeads followed by sample prep for TEM. Using stereology on the ultrathin sections of Dynabeads coated with exosomes can give you exosome counts per bead. Previously we compared the exosome numbers counted by stereology with the signal we obtained by flow cytometry. Increased counts by TEM correlated with increased signal by flow as well as western blot.

How can exosomes having different origins be differentiated if the sample is serum?

In terms of exosome characterization and comparison, this can be performed as we did it in our publication (Oksvold et al., Clinical Therapeutics 2014 June 36, 847-862.e1, PMID:24952935). We characterized 4 different B-cell lymphoma derived exosomes by starting to compare the expression of typical exosomal markers, such as tetraspanins and B-cell specific markers. Characterization was then performed after pre-enrichment of the exosomes and again after isolating exosomes with Dynabeads magnetic beads that target CD81+ and CD63+ exosomes. Interestingly, the expression of both exosomal markers and B-cell markers differed between all 4 B-cell lymphoma derived exosomes. The complex nature of serum as a starting sample will add to the complexity. Here a pre-clearing step (e.g., using size-exclusion chromatograph prior to Dynabeads isolation) is recommended.

For FACS analysis with Dynabeads magnetic beads, which capture antibodies (CD9, CD63, CD81...) work the best for detecting exosomes pre-enriched from plasma?

Currently, we do not have an optimized protocol for exosome isolation from plasma using Dynabeads. However, we have been able to capture CD9 positive exosomes from plasma using “Dynabeads CD9” [i.e., Exosome Human CD9 Isolation Reagent (from Cell Culture), Cat. No. 106-14D], after performing a size-exclusion chromatography. For testing, we recommend these magnetic beads, which works very well in all samples tested given that the exosomes are CD9 positive.

What is the recommended volume used for purifying exosomes from pre-enriched exosomes using ultracentrifugation, and how high is your yield (e.g., total amount of protein)? How does this compare to the direct method using Dynabeads magnetic beads?

What we demonstrated in the webinar was a comparison between isolation of pre-enriched exosomes using Dynabeads with a direct capture strategy using Dynabeads. In order to compare it was critical to keep the binding kinetics constant and adjust for the exosome concentration factors. In our experiments, the concentration factors were x42 for exosomes pre-enriched by ultracentrifugation and x16 for exosomes pre-enriched by precipitation. We added equal amounts of exosomes to all isolations, keeping the amount of isolation beads at 20 µL and the isolation volume at 100 µL. In theory, we would expect to capture the same amount of exosomes in all three isolations. However, we seemed to lose vesicles when performing pre-enrichment by ultracentrifugation—which is not surprising. The idea that ultracentrifugation is the ultimate method to capture all vesicles seems to me to be a bit strange, as it requires very skilled technicians to avoid vesicle loss during preparation. When there is enough exosome in the samples, direct capture is the better option. I recommend performing pre-enrichment followed by capture using Dynabeads only when the starting sample contains few exosomes or the volume is large.

Which concentration of Dynabeads magnetic beads we use depends on the downstream application. Typically, for flow cytometry, we want as many exosomes as possible per bead to provide a strong signal. Therefore, few beads are used. For downstream western blotting, many beads are used to allow maximum number of exosomes to dock to a large surface area.

For flow cytometry, we use 20 µL of 1x107 Dynabeads per 100 µL isolation volume

For western blotting, we use 20 µL of 1.3x108 Dynabeads per 100 µL isolation volume

How much protein do I start with?

I would not recommend paying too much attention to the protein concentration as an estimation for the amount of exosomes in your preparation. Our experience is that it is not always a good correlation. We have mostly been working with cell culture media and urine as starting samples, and even in such “simple” solutions (at least compared to plasma/serum) the correlation is not that good. That’s why we started to pay attention to the cell culture growth conditions and standardized the exosome harvest conditions in order to harvest at the correct time and increase the reproducibility in our system. It’s also the reason why we use Dynabeads magnetic beads to collect exosomes during cell growth as a much better method to estimate the amount of exosomes. I would assume that there will be even less correlation between protein concentration and exosome content in plasma and serum.

How much/many Dynabeads magnetic beads should I use for exosome isolation?

The amount of beads used to capture exosomes will depend on the downstream application. For flow cytometry you will need to use as few beads as possible in order to obtain a strong signal, i.e., to maximize the load of exosomes per bead and provide signal when each bead is traveling through the flow cell. This is very different for methods such as western blotting which will require more total exosomes to get a signal. This can be done by increasing the amount of beads significantly during isolation in order to provide a larger total surface area.

# beads used for western blotting: 20 µL of beads (stock solution of 1.3x108 beads/mL) in 100 µL of volume containing your exosomes

# beads used for proteomics: If this refers to mass spectrometry applications, we currently have not optimized a protocol for that. However, I will assume that the conditions should look more like the conditions for western blotting rather than flow cytometry in order to capture more exosomes for the analysis.

How can exosomes be released from Dynabeads magnetic beads?

Currently we have not optimized a release method to get exosomes off the beads. Low pH or high salt have been used to release protein complexes from the beads, and I know that some researchers also use these techniques to release exosomes. However, I have not seen any data about the functionality of the exosomes after such treatment. If the downstream application is mass spectrometry, the exosomes can be processed for MS in the presence of the beads.

Which markers can be used for exosome characterization?

As far as I know (even after consulting with many key opinion leaders in the field of extracellular vesicles), there is currently no consensus about a general exosome marker that will be found on all exosomes. The current recommendation from the research community is to combine detection of many membrane-bound or membrane-associated proteins in order to verify the presence of membranes. In our own experience, targets such as the CD63 and CD81 are found in many different exosome preparations, as is CD9. However, we could detect at least 2 different cell lines releasing exosomes which were CD9 negative (Jurkat cells and several B-cell lymphoma cells). In house, we usually label for CD9, CD81 and CD63 to verify the presence of membranes in our preparation using the following detection antibodies:

These detection antibodies have been validated for use with both pre-enriched exosomes and those isolated with Dynabeads magnetic beads. Tsg101 and annexins have also been mentioned as useful targets. It is also important to characterize the host cell in terms of different markers to determine the level of contaminating vesicles from these organelles:

ER: HSP90B1, calnexin

Golgi: GM130

Mitochondria: cytochrome C

Nucleus: Histones

How can exosomes be stored?

We store exosomes in PBS with 0.1% BSA, as this is the buffer used to dissolve our pellet after precipitation. The isolation efficiency is not changed after freezing at -80°C compared to freshly made exosomes. For direct isolation from cell culture media or even urine we have frozen the cell culture without adding any form of cryo-protectant such as glycerol.

Do you have Dynabeads products for isolating exosomes derived from mouse cells?

Nanoparticle tracking analysis (NTA) is probably the most-used method for obtaining information about the average size and concentration of the particles in the solution. However, this method cannot distinguish between vesicles and protein aggregates of the same size. Therefore, we recommend combining this method with microscopic techniques in order to verify the presence of membranes. We have paid a lot of attention to the cell culture and exosome harvest conditions in order to produce equal amounts of exosomes per harvest; this is far more important in our opinion than estimating the protein concentration of the preparation.

The way Dynabeads magnetic beads are designed and produced for cell isolation, protein isolation, nucleic isolation, T-cell activation and expansion (and now also exosome capture and isolation) helps ensure a high degree of reproducibility. All the beads are the same size and have a surface chemistry that provides very low unspecific binding and high signal/noise ratio. With this in mind, isolating exosomes using Dynabeads (paying attention to the bead concentration and capture volume) will enable constant binding kinetics. This setup will not give you the exact number of exosomes on the bead surface, but it will provide increased signal as a result of increased input in a linear way. Therefore, the beads can be used to measure exosomes, e.g., to monitor the exosome release efficiency from cells.

Electron microscopy must be used to estimate the number of exosomes per bead, i.e., isolate exosomes with Dynabeads followed by sample prep for TEM. Using stereology on the ultrathin sections of Dynabeads coated with exosomes can give you exosome counts per bead. Previously we compared the exosome numbers counted by stereology with the signal we obtained by flow cytometry. Increased counts by TEM correlated with increased signal by flow as well as western blot.

How can I deplete the exosomes from the cell culture media?

One thing to consider is using Dynabeads products as depletion beads. For example, use Dynabeads CD9 (Cat. No. 10614D), CD81 (Cat. No. 10616D) or CD63 (Cat. No. 10606D) reagents in a mixture or even sequentially to deplete exosomes from the solution.

Can exosomes be recovered from viral particles? If so, how?

Separation of exosomes from viral particles could be done in two ways – either by negative isolation or positive isolation. If antibodies are available for targeting viral antigens these can be coupled to Dynabeads magnetic beads, either non-covalently to ligand beads such as Dynabeads Protein G (Cat. No. 10003D) or covalently to surface-activated beads such as Dynabeads M-270 Epoxy (Cat. No. 14302D) or the Dynabeads Antibody Coupling Kit (Cat. No. 14311D). In this way, one can deplete the sample of viral particles and leave the exosomes untouched. Alternatively, the ready-made Dynabeads CD9 (Cat. No. 10614D), CD81 (Cat. No. 10616D) or CD63 (Cat. No. 10606D) could be used to pull out the exosomes from the solution.

Can the isolation kits be used for mass spectrometry experiments downstream?

Our experience (and others as well) is that there is little correlation between the protein measurement and the number of exosomes in the preparation. As far as I know there is no real consensus regarding which method to use. Many researchers use nanoparticle tracking analysis (NTA) to measure number of vesicles and size distribution, but this methods does not give information about any contaminating vesicles in the same size range. Therefore, electron microscopy should always be combined with this method. One way of checking the presence of contaminating vesicles might be to perform a western blot to detect a protein such as gp96, an ER-related protein.

We have focused on standardizing cell culture and exosome harvest conditions to increase the chance of loading equal amounts of each exosome preparation. In addition, it might be possible to set up a standard curve using different amount of protein input, perform a western blot and relate the signal of target to these bands on the blot. In addition, as we presented in the webinar, we also pay attention to the amount of exosome input (and the concentration factor if we start with pre-enriched material), isolation volume and also the amount of capture beads in order to keep binding kinetics and the exosome input constant so that we can compare results between different samples.

Yes. We typically use fresh media, or one stored at 4°C for up to a week, but the exosomes frozen at -20°C or -80°C work fine. Ideally the exosomes should be frosted/defrosted only 2 to 3 times; multiple freeze/thaw cycles partially damage exosomes/EV.

Does Thermo Fisher Scientific offer a system for making exosomes in vitro for transfection for incorporation of RNA or proteins?

Thermo Fisher Scientific offers an arsenal of transfection reagents, including the widely used Lipofectamine 2000 and RNAi MAX transfection reagents. Exosomes are definitely promising from the delivery standpoint. However, there are many challenges and limitations in using complex biological vesicles for delivery purposes.

What kind whole genome amplification (WGA) kit can be used to increase the DNA concentration from tumor-derived exosomes?

Exosomal DNA is not well studied. It is crucial to obtain very clean exosomal preparations, and to characterize them initially by all available means. In the very least, use NanoSight nanoparticle tracking analysis (NTA) to ensure that your protocol is not recovering large microvesicles along with exosomes to avoid substantial contamination with long DNA. I suggest using a large sample volume to recover enough exosomes and their DNA/RNA cargo. In general, it is a very interesting area of exploration. I recommend using a large sample volume to recover enough exosomes and their DNA/RNA cargo. Manipulate with at least 4 mL of serum/plasma to recover enough nucleic acid to run Bioanalyzer, NanoDrop instruments and other typical means of analysis, prior to sequencing.

If most miRNAs are attached to proteins and only 20% is in exosomes, do the methodologies described today provide enough separation to distinguish the two sources of microRNAs?

A substantial part of miRNA is associated with proteins. Ago2 and other protein-miRNA complexes can be isolated with antibodies coupled to Dynabeads magnetic beads. Exosomes can be precipitated from the same sample with the Total Exosome Isolation Reagent (Cat. No. 4478359) afterwards, or exosomal populations can be isolated with Dynabeads CD9, CD63 or CD81 products. We are not aware of any papers that discuss separately recovering different entities from the same body fluid source (protein-miRNA, exosomes, microvesicles, etc.).

Have you compared the yield of exosomal RNA between direct capture and pre-enrichment protocols?

With the Dynabeads magnetic beads, the typical initial step is pre-enrichment with the Total Exosome Isolation Reagent (Cat. No. 4478359) (e.g., you recover total exosomal population from cell media, then isolate exosomes positive for CD9, CD63 or CD81 with Dynabeads). As now demonstrated, exosomes can also be captured directly using Dynabeads products.

According to the literature it seems that 18S mRNA is absent from exosomes. Can you comment on that?

We have analyzed exosomal RNA content by sequencing and qPCR, and these vesicles definitely include some 18s RNA. In our opinion, the early papers were misleading. One team claimed that exosomal cargo is miRNA- and mRNA-negative, and everyone else accepted this as a dogma. In reality, exosomes contain ribosomal RNA, tRNA, many non-coding RNAs, and a large fraction of miRNA is actually associated with proteins (e.g., Ago 2) not exosomes.

Could you describe the workflow for exosomal RNA isolation and how you perform the PCR?

Details regarding exosomal isolation and PCR can be found in the follow articles:

What controls should be included to ensure maximal signal/noise ratio for miRNA isolation only from exosomes?

I did not see any papers yet on the topic of separately recovering different entities from the same body fluid source (protein-miRNA, exosomes, microvesicles, etc). However, know that a number of teams are looking into fractions vs. analysis of whole serum or plasma RNA.

What is the recovery rate of exosomes with magnetic beads and what was the working volume of the cell sample you used?

The concentration of Dynabeads magnetic beads we use depends on the downstream application. Typically, for flow cytometry, we want as many exosomes as possible per bead to provide a strong signal. Therefore, few beads are used. For downstream western blotting, many beads are used to allow maximum number of exosomes to dock to a large surface area.

For flow cytometry, we used 20 µL of 1x107 Dynabeads per 100 µL isolation volume

For western blotting, we used 20 µL of 1.3x108 Dynabeads per 100 µL isolation volume

The figures below show the effect of bead amount on signal/noise (S/N) in flow cytometry (Panel A) and western blotting (Panel B). Here “1x” beads represent the amount we use for flow cytometry and “25x” represent the amount use for western blotting.

That is a very interesting question. I know that Hendrix et al compared ultracentrifugation (UC) methods with precipitation methods, claiming UC with gradients were superior. Also Tauro et al (2012) performed comparison where immune-capture with beads was considered to be the best method. I guess this reflects the fact that there is no consensus yet. In my opinion, the fewer steps involved in the preparation, the better the result. Clearly, performing UC as a pre-enrichment step will result in loss of vesicles—even if you are a really skilled operator. So even though the purity of vesicles is high, there is no control over which vesicles are lost during processing.

Has anyone had success visualizing the exosomal pellets produced following centrifugation with the total exosome isolation reagent? We would like to quantify the exosomes in our culture media following different cell treatments, and I find it hard to be confident that I am not losing any exosomal material during my media removal/pellet re-suspension.

What we know from in-house experience is that there is a risk of losing material during preparation—as we demonstrated in the webinar where material seems to be lost during ultracentrifugation (UC). The amount lost will of course also depend on the experience of the operator. The loss of material seems to be less when performing precipitation as a way of pre-enriching exosomes. This protocol is much more simple compared to an ultracentrifugation protocol. However, by omitting this step in the workflow and using Dynabeads magnetic beads to capture exosomes this can be used as a measurement of the exosome release during cell culture in order to, e.g., harvest at the best time point. Of course, this approach might lose some exosomes if they lack the marker used for isolation. But the depletion data shows that using enough beads for isolation results in a high degree of depletion. So, in conclusion, I strongly believe that currently there is no method out there which will ensure isolation of absolutely all exosomes in the preparation. So currently the best way of characterization is to combine several methods.

Is it better using glycerol (in which case, what concentration?) to perform better cryogenic storage or is it sufficient to freeze exosomes in the medium in which they have been isolated?

In terms of storing the exosomes, we use PBS w/ 0.1% BSA as this is the buffer used to dissolve the pellet after precipitation. The isolation efficiency is not changed after storing at -80°C compared to freshly made exosomes. For direct isolation from cell culture media or even urine we have frozen preparations this way without adding any form of cryo-protectant such as glycerol. We have not tested the use of glycerol so I do not know how this will affect the isolation efficiency.

How does the Total Exosome RNA and Protein Isolation Kit work?

The Total Exosome RNA and Protein Isolation Kit is developed specifically for exosome samples, and uses an initial Acid-Phenol: Chloroform extraction. This is followed by addition of ethanol to the aqueous phase, and then the sample is passed through a glass-fiber containing filter cartridge to immobilize the RNA. The filter is washed, and the RNA is eluted with a low ionic-strength solution. The kit recovers all RNA longer than ~10 nt up to several kb. (The majority of exosomal RNA ranges in size from ~20–300 nt.) The product manual also describes an additional protocol for enrichment of short RNA (<200 nt), but we recommend the total RNA isolation protocol in order to maximize recovery of all RNA, including various mRNA, rRNA, and ncRNA fragments.

The kit also provides an option to recover protein from the same sample through the use of the Exosome Resuspension Solution. The kit can be used to isolate RNA and protein from exosomes purified from any sample type using either the Total Exosome Isolation reagents or any other protocol such as ultracentrifugation.

When ethanol is added, the solution turns turbid. Does this affect efficiency of RNA recovery?

No, the described effect does not have a negative effect on the RNA recovery.

How much RNA can be recovered from the exosomes?

This can vary depending on the sample type, volume of sample, isolation method, and exosome content/concentration. Listed below are some examples:

In both cases, these amounts of RNA are sufficient for RNA library prep for Ion Torrent PGM or Proton sequencing. For real-time PCR analysis, substantially smaller amounts of RNA are needed and much lower sample volumes can be utilized. For example, RNA recovered from 3 μL serum or 30 μL cell media is enough for one RT-qPCR reaction.

What is the current method for exosome isolation?

Traditional isolation of exosomes from cell culture media and body fluids is a tedious and difficult process, typically based on ultracentrifugation in combination with sucrose density gradients or sucrose cushions to float the relatively low-density exosomes away from other vesicles and particles. These protocols can take up to 30 hours, require an ultracentrifuge and extensive training. Despite these drawbacks, this is still the most typical approach for exosome isolation.

However, within the last couple of years, several reagents for isolating exosomes from cell culture media and various body fluids have been made commercially available from Life Technologies and other companies. The Total Exosome Isolation reagents available from Life Technologies allow for the recovery of exosomes, using a very short and reliable protocol. This approach is becoming more and more popular.

Which products for exosome research are available from Life Technologies?

A number of products developed specifically for exosomes research are available, as described here: www.lifetechnologies.com/exosomes. We are continuously expanding this product portfolio.

In addition, we offer many reagents, kits and instruments for analysis of exosomal cargo including: RT-qPCR with TaqMan assays, next generation sequencing on Ion Torrent PGM, Proton and SOLID for RNA and Western blotting instruments and reagents for protein research.

How do the Total Exosome Isolation reagents work, and what makes them different from the other techniques for exosome isolation?

In the course of development of reagents for isolation of exosomes we evaluated many different technologies (e.g. ultracentrifugation, ultrafiltration, gel-filtration columns, HPLC, and filters). We also evaluated more advanced approaches (e.g. precipitation using various polymers, and bead and column binding using antibodies and various lectins). In addition, we evaluated commercially available products from different companies.

We then selected one of the polymers, based on its superior performance, which became the key component of the Total Exosome Isolation reagents (Patent application filed). By tying up water molecules, the reagent forces less-soluble components, such as vesicles, out of solution which allows them to be collected by a short, low-speed centrifugation. The recovered exosomes are then ready for either biological studies or end-point analysis. All Total Exosome Isolation reagents share the same core compound, but have each been carefully optimized (incl. adjustments in protocols), to enable efficient isolation of exosomes from specific sample types.

Do the Total Exosome Isolation reagents allow isolation of exosomes from other species?

Yes, the reagents can also be used with mouse samples. The reagents can potentially also isolate exosomes from samples of any species, but have so far only been tested with human and mouse.

Does the Total Exosome Isolation (from serum) reagent work on plasma samples?

Plasma is a more challenging type of sample compared to serum, as it has high levels of clotting factors. The current serum reagent will work on plasma, but the preparation will likely contain more contaminating proteins and microvesicles. For plasma, we recommend using the Total Exosome Isolation (from plasma) reagent, with a protocol specifically optimized to handle plasma.

hat is the purity of the exosomes recovered with the Total Exosome Isolation reagent, is it similar to ultracentrifugation?

The obtained sample contains all exosomes, with insignificant amounts of some additional microvesicles and large protein molecules/complexes that have been co-precipitated (in case of serum and some of the other body fluids). This purity level works fine for most applications, and is balanced by the method benefits. The benefits include: 1) A fast and simple workflow, 2) No need for special equipment (such as an ultracentrifuge), 3) Complete recovery of exosomes, 4) The flexibility to work with small sample volumes (e.g. 100 μL), 4) The capability to process multiple samples in one experiment.

To allow for the recovery of a very clean population of exosomes following an initial purification with the Total Exosome Isolation (from cell media) reagent, we also offer Dynabeads coupled with anti-CD63 antibodies. This additional step will increase workflow time, and the final yield of the exosomes will be lower - but for projects that require an ultra-clean population of CD63-positive exosomes from cell culture media this is the best option. Streptavidin-coupled Dynabeads are also available for use with your choice of biotinylated antibody - specific for a specific exosome sub-population.

These products can be used not only for isolation of highly pure exosome sub-populations, they also allow detection of exosomes with flow cytometry—something that has otherwise been extremely difficult to achieve due to their small size. We believe that these products are currently the best options for exosome isolation.

How do the Total Exosome Isolation reagents compare to the ExoQuick reagents from System Biosciences?

The Total Exosome Isolation reagents have a unique formulation (patent application filed). Carefully optimized reagents allow recovery of exosomes from major body fluids and cell media. The yield and purity of the exosomes recovered with the products supplied by us is equivalent—and sometimes improved—compared to ExoQuick reagents. In addition, our reagents enable isolation of the total exosome population from more sample types, and the products are part of a larger, complete workflow solution from isolation to characterization to in vivo studies.

What is the minimal volume of starting sample I can use to isolate exosomes?

For each reagent, the minimal volume tested can be found listed in the product manual. For most body fluids the minimum volume tested is 100–200 μL, and slightly larger for urine (800 μL) and cell culture media (1 mL). Smaller volumes can be used—especially for serum and plasma—but we’ve found that the minimums listed here provide a usable amount of exosomes for multiple downstream applications.

If I recover exosomes with the Total Exosome Isolation reagent from a larger sample volume, >5 mL, will I need to spin them for a longer time?

For larger sample volumes than those recommended in the manuals, a longer centrifugation is recommended to ensure maximum recovery of the exosomes. The exact time will depend on several factors including: the rotor, the sedimentation coefficient of the exosomes, size of the tube, sample volume and type, and centrifugation speed. For example, for 5-10 mL sample volumes, 1 hour of centrifugation at 10,000 x g is sufficient to pellet the exosomes. But if the sample volume is larger, or centrifuge cannot generate 10,000 x g—longer centrifugation times should be used.

How many exosomes can be isolated from cell media and serum?

The numbers will be somewhat different for different cell lines, and depending on whether you add exosome-depleted FBS or no FBS or synthetic media. Here’s an example: HeLa cells grown to ~ 2 x 107 per T175 flask in 30 mL cell culture media, with exosome-depleted FBS. From 1 mL of this cell media, one can recover ~4-8 x 109 exosomes with the Total Exosome Isolation (from cell culture media) reagent, as measured with NanoSight LM10. From 100 μL serum you can recover ~1.5-3 x 1011 exosomes—with the Total Exosome Isolation (from serum) reagent.

There are two protocol options for plasma samples, which one should I choose?

Unlike serum, plasma contains numerous clotting factors and some additional protein which can make the sample difficult to handle. To ease this difficulty , we’ve provided two protocol options: One with Proteinase K treatment , and one without. The protocol including Proteinase K treatment is most useful when the end goal is analysis of exosomal RNA or protein cargo. This protocol can also be used to isolate exosomes for use in other downstream applications, but is most useful for RNA and protein analysis. The protocol without Proteinase K treatment also isolates good quality exosomes, just not quite as pure as the protocol with Proteinase K treatment. The protocol without Proteinase K treatment is more useful when isolating exosomes that will be used for surface protein analysis or electron microscopy identification.

Due to time constraints, I cannot finish the protocol in a single day. Can I precipitate my exosomes overnight?

Is there any reagent left in the exosome preparation after isolation? If so, can this interfere with the downstream biological studies?

The reagent does not bind to the surface of the exosomes, and only trace amounts remain in the exosome pellet after isolation. This should not interfere with downstream biological studies. However, it is important to completely remove the supernatant prior to resuspending the exosome pellet in PBS or other buffer of choice. If you still have concerns about trace amounts of the reagent, this can be removed by dialysis or using the Exosome Spin Columns (MW 3000).

Is it OK if the exosome pellet is not always visible after isolation? Can we leave some supernatant in the tube, just to be sure not to lose any of the exosome pellet?

Serum and plasma contain a very high number of exosomes, thus the pellet is visible even if you isolate exosomes from as little as 100 μL. Other body fluids, such as urine, or cell culture media, have significantly lower concentration of exosomes, and the pellet is often not visible after centrifugation. Since the pellet sticks very tightly to the tube, it is OK to remove the supernatant completely prior to resuspending in PBS. If needed, it is easy to label the tube so you know where the pellet will adhere upon centrifugation. It is absolutely crucial to remove the supernatant completely. If you don’t, there will be a significant amount of the reagent left, and when you resuspend the exosomes some of them might be still in the form of aggregates, at the bottom of the tube.

How are exosomes defined?

The current definition of exosomes is complex, with no absolute consensus in the field. Typically, exosomes are defined as vesicles floating in sucrose solutions at a density ~1.13 to 1.19 g/ml during ultracentrifugation-based isolation, with an expected size of 30–150 nm (based on electron microscopy analysis). Exosomes can also be defined and identified by their surface protein markers, which include: tetraspanins (CD63, CD81, CD9) and others like ALIX. Currently, there are no appropriate tools nor sufficient knowledge in the field to set a clear and simple definition of exosomes that would differentiate them from other micro-/nanovesicles.

What are exosomes composed of?

Exosomes are tiny vesicles (30–150 nm) containing protein and/or RNA cargo, within a lipid bi-layer membrane. Exosomes can differ extensively in both their cargo and surface proteins, and different cell types can secrete different —sometimes multiple—types of exosomes.

What is the mechanism of exosome formation?

Exosomes are classically described as vesicles originating from the endocytic pathway through fusion of multivesicular bodies with the plasma membrane. They are a part of a larger family of vesicles secreted by cells—including microvesicles, ectosomes and shed particles–which originate by direct budding from the plasma membrane. It is extremely challenging to separate these entities using currently available techniques and instruments, due to overlap in their size, density and overall composition similarities.

How do I know that I’ve isolated exosomes, and not other types of vesicles?

In addition to their 30–150 nm size, to be categorized as exosomes, the vesicles should be positive for certain surface protein markers such as tetraspanins. The most widely accepted marker is CD63, but CD81, CD9 are utilized as well. Western blotting for these targets on the sample of interest is a relatively simple way to confirm that the vesicles are indeed exosomes. However, the current definition of exosomes is not set in stone, as there is no absolute consensus in the field. It will probably take another several years to come up with the exact specification and nomenclature for all nano-/microvesicles (including exosomes).

How are exosomes visualized and/or counted?

Exosomes are too small (30–150 nm) to be seen using a regular microscope, as this is limited to objects that are at least several micrometers in size. Structures as small as bacteria, and even very large viruses such as vaccinia virus (approx. 0.5µm in size), can be observed by light microscopy. Yet the resolution of the light microscope is too low for observing exosomes.

The typical methods of analysis for exosome size distribution include the NanoSight instrument and electron microscopy. Although very different in methodology, both technologies allow you to study nanoparticles down to 10 nanometers in size.

To determine the concentration of exosomes in the sample, you can use the NanoSight instrument or the Izon instrument. The NanoSight instrument enables counting and sizing of nanoparticles (10–1000 nm) using light scattering and browning motion, while the Izon instrument accomplishes the same thing using nanopore analysis.

What is the best way to store my exosomes?

For short-term, exosomes can be stored at 4°C for up to 1 week. For long-term storage, exosomes can be stored at -20°C or -80°C. When storing exosomes long-term, it is important to consider if they will need to be thawed more than once. If multiple applications (and thus multiple thaws) is required for analysis, we recommend aliquoting the exosome suspensions into multiple tubes so that each tube will only undergo one freeze/thaw cycle. We have found that multiple freeze thaw cycles can cause damage to the exosomes and reduce their numbers.

What should I do when my Westerns do not seem to work?

There are several typical reasons for this:

Not enough sample volume added. Exosomes can contain a fairly low amount of protein cargo. For an initial experiment, we recommend adding as much of the sample as possible

Antibodies are not optimal. We suggest testing antibodies (e.g. anti-CD63 or other exosomal marker) from 2–3 manufacturers, carefully checking what concentration is recommended. Also, they should ideally be used fresh, and need to be stored properly.

Depending on the exosomal surface marker, certain gel conditions might be more optimal for the target antibody (e.g. reducing/non-reducing and denaturing/non-denaturing). We suggest checking with the manufacturer and exosome community about which conditions are recommended for the specific marker you are targeting and the specific antibody you are using.

General Western techniques. Westerns can be tricky, and we recommend the use of a positive control for initial testing to make sure the entire workflow is functioning as it should. Any protein or antibody can be used as long as they meet the conditions you need (e.g. denaturing vs. non-denaturing). In addition, when picking the protein, try to steer clear of those that are present at very high or very low concentrations in your sample to prevent overloading the blot or total absence of signal.